Submitted 12 January 2014Accepted 10 March 2014Published 25 March 2014

Corresponding authorElizabeth J. Harry,liz.harry@uts.edu.au

Academic editorSiouxsie Wiles

Additional Information andDeclarations can be found onpage 20

DOI 10.7717/peerj.326

Copyright2014 Lu et al.

Distributed underCreative Commons CC-BY 3.0

OPEN ACCESS

Manuka-type honeys can eradicatebiofilms produced by Staphylococcusaureus strains with differentbiofilm-forming abilitiesJing Lu1, Lynne Turnbull1, Catherine M. Burke1, Michael Liu1,Dee A. Carter2, Ralf C. Schlothauer3, Cynthia B. Whitchurch1 andElizabeth J. Harry1

1 The ithree institute, University of Technology Sydney, Sydney, NSW, Australia2 School of Molecular Bioscience, University of Sydney, Sydney, NSW, Australia3 Comvita NZ Limited, Te Puke, New Zealand

ABSTRACTChronic wounds are a major global health problem. Their management is difficultand costly, and the development of antibiotic resistance by both planktonic andbiofilm-associated bacteria necessitates the use of alternative wound treatments.Honey is now being revisited as an alternative treatment due to its broad-spectrumantibacterial activity and the inability of bacteria to develop resistance to it. Manyprevious antibacterial studies have used honeys that are not well characterized, evenin terms of quantifying the levels of the major antibacterial components present,making it difficult to build an evidence base for the efficacy of honey as an antibiofilmagent in chronic wound treatment. Here we show that a range of well-characterizedNew Zealand manuka-type honeys, in which two principle antibacterial compo-nents, methylglyoxal and hydrogen peroxide, were quantified, can eradicate biofilmsof a range of Staphylococcus aureus strains that differ widely in their biofilm-formingabilities. Using crystal violet and viability assays, along with confocal laser scanningimaging, we demonstrate that in all S. aureus strains, including methicillin-resistantstrains, the manuka-type honeys showed significantly higher anti-biofilm activitythan clover honey and an isotonic sugar solution. We observed higher anti-biofilmactivity as the proportion of manuka-derived honey, and thus methylglyoxal, in ahoney blend increased. However, methylglyoxal on its own, or with sugar, was notable to effectively eradicate S. aureus biofilms. We also demonstrate that honey wasable to penetrate through the biofilm matrix and kill the embedded cells in somecases. As has been reported for antibiotics, sub-inhibitory concentrations of honeyimproved biofilm formation by some S. aureus strains, however, biofilm cell suspen-sions recovered after honey treatment did not develop resistance towards manuka-type honeys. New Zealand manuka-type honeys, at the concentrations they can beapplied in wound dressings are highly active in both preventing S. aureus biofilmformation and in their eradication, and do not result in bacteria becoming resistant.Methylglyoxal requires other components in manuka-type honeys for this anti-biofilm activity. Our findings support the use of well-defined manuka-type honeys asa topical anti-biofilm treatment for the effective management of wound healing.

How to cite this article Lu et al. (2014), Manuka-type honeys can eradicate biofilms produced by Staphylococcus aureus strains withdifferent biofilm-forming abilities. PeerJ 2:e326; DOI 10.7717/peerj.326

Subjects Microbiology, Infectious DiseasesKeywords Staphylococcus aureus, Biofilm, Honey, Antibacterial, Wounds, Methylglyoxal

INTRODUCTIONChronic wounds currently affect 6.5 million people in the US. These wounds are difficult

to treat and estimated to cost in excess of US $25 billion annually, with significant increases

expected in the future (Sen et al., 2009). A wound is generally considered chronic if it has

not started to heal by four weeks or has not completely healed within eight weeks (McCarty

et al., 2012). Such prolonged, non-healing wounds are caused by a variety of factors, with

bacterial infection being a significant contributor.

In chronic wounds, as with everywhere on earth, bacterial cells predominantly exist

as biofilms, where cells are embedded within a matrix of polysaccharides and other

components. This matrix affords resistance to environmental stresses such as altered pH,

osmolarity, and nutrient limitation (Fux et al., 2005). The matrix also limits access of

antibiotics to the biofilm embedded cells (Ranall et al., 2012), which are up to 1,000 times

more recalcitrant to these compounds than planktonic cells (Hoyle & Costerton, 1991).

Planktonic bacteria may also contribute to pathogenesis, as their release from biofilms has

been proposed to maintain the inflammatory response within the wound (Ngo, Vickery

& Deva, 2012; Wolcott, Rhoads & Dowd, 2008), as well as allowing seeding to other areas

(Battin et al., 2007; Costerton et al., 2003). Along with the difficulties of treating biofilm in-

fections, the emergence of resistance to multiple antibiotics has exacerbated the problem of

chronic wound treatment (Engemann et al., 2003; Projan & Youngman, 2002). Thus, there

is an increasing need for new approaches to combat bacterial biofilms in chronic wounds.

Honey has been used to treat acute and chronic wound infections since 2500 BC

(Forrest, 1982; Molan, 1999; Simon et al., 2009). Honey possesses a number of antimicrobial

properties including high sugar content, low pH, and the generation of hydrogen peroxide

by the bee-derived enzyme glucose oxidase (Stephens et al., 2009). However, not all honeys

are the same and their antimicrobial properties vary with floral source, geographic

location, weather conditions, storage (time and temperature) and various treatments,

such as heat (Al-Waili et al., 2013; Allen, Molan & Reid, 1991; Molan, 1999; Sherlock et al.,

2010). These factors lead to differences in the levels of various antibacterial components.

Manuka honey is derived from Leptospermum scoparium bush and is particularly potent

(Adams et al., 2008; Allen, Molan & Reid, 1991; Kwakman et al., 2011). This is believed to be

largely due to the high levels of the reactive dicarbonyl methylglyoxal (MGO) (Adams et al.,

2008; Mavric et al., 2008), which is highly inhibitory to bacterial growth (Lu et al., 2013).

Other antimicrobial compounds in honeys include bee defensin-1 (Kwakman et al., 2010;

Kwakman & Zaat, 2012), various phenolic compounds and complex carbohydrates (Adams

et al., 2008; Gresley et al., 2012; Mavric et al., 2008; Molan, 1999; Weston, Brocklebank & Lu,

2000). The combination of these diverse assaults may account for the inability of bacteria to

develop resistance to honey (Blair et al., 2009; Cooper et al., 2010), in contrast to the rapid

induction of resistance observed with conventional single-component antibiotics (Colsky,

Kirsner & Kerdel, 1998; Cooper, 2008).

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 2/25

A few studies have examined the effect of manuka honey on biofilms, showing it

to be active against a range of bacteria, including Group A Streptococcus pyogenes,

Streptococcus mutans, Proteus mirabilis, Pseudomonas aeruginosa, Enterobacter cloacae

and Staphylococcus aureus (Alandejani et al., 2008; Maddocks et al., 2013; Maddocks et

al., 2012; Majtan et al., 2013). However, the levels of reported anti-biofilm activity are not

consistent among these studies. This is highly likely to be at least in part due to differences

in the levels of the principle antibacterial components in the honey, MGO and hydrogen

peroxide, which varies with the floral and geographic source of nectar, the honey storage

time and conditions, and any possible other treatments that may have occurred. All these

conditions affect the antimicrobial activity of honey (Adams et al., 2008; Al-Waili, Salom

& Al-Ghamdi, 2011; Sherlock et al., 2010; Stephens et al., 2009), but are often not reported.

Importantly, medical-grade honeys, while often composed primarily of manuka, can

also contain honey derived from other flora sources, which can alter the levels of various

antimicrobial components. Therefore, it is imperative to use well-characterized honeys

to enable both accurate comparisons among studies, and the rigorous assessment of the

potential of medical-grade honey to be used in wound treatment in the clinic.

Here we have performed biomass and viability assays, as well as confocal scanning light

microscopy to examine the anti-biofilm activity of four NZ manuka-type honeys, clover

honey and an isotonic sugar solution on a range of S. aureus strains that differ widely in

their biofilm-forming ability. These honeys have been well characterized in terms of their

geography, floral source and the level of the two principal antibacterial components found

in honey, MGO and hydrogen peroxide. We demonstrate that the manuka-type honeys

are highly active in both the prevention and elimination of methicillin-sensitive and

methicillin-resistant S. aureus biofilms. The antibiofilm activity was highest in the honey

blend that contained the highest level of manuka-derived honey; although the same level

of MGO, with or without sugar, could not eradicate biofilms. This suggests that additional

factors in these manuka-type honeys are responsible for their potent anti-biofilm activity;

and emphasise the importance of characterizing honey in order to understand and choose

the best honey product for wound management.

MATERIALS AND METHODSHoney samplesThe New Zealand (NZ) honey samples used in this study are listed in Table 1, and

include monofloral manuka honey, Medihoney (a manuka-based medical grade honey;

Comvita NZ Ltd), a manuka/kanuka blend, and clover honey (a white New Zealand

honey). All honey samples were supplied by Comvita NZ Ltd. (Te Puke, New Zealand).

The harvesting and geographic information for these honeys, as well as the levels of the

three major antimicrobial components: methylglyoxal (MGO), di-hydroxyacetone (DHA)

and hydrogen peroxide, are listed in Table 1. All samples were stored in the dark at 4 ◦C and

were freshly diluted in Tryptone Soya Broth (TSB) immediately before use in assays. All

honey concentrations are expressed as % w/v.

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Table 1 Harvesting and chemical information for the tested NZ honey samples.

Honey type Harvest period Area Floral source Major antimicrobial composition

DHAa MGOa H2O2b

Manuka Spring 2010 Hokianga, Northland, NZ Leptospermum scoparium var.incanum

4277 958 0.34

Medihoney Spring 2010 Northland, NZ Leptospermum scoparium var.incanum + Kunzea ericoides

883 776 0.31

Manuka/kanuka Summer 2010/11 Hokianga, Northland, NZ Leptospermum scoparium var.incanum + Kunzea ericoides

652 161 0.68

Clover N/A* Balcutha, Otago, NZ Trifolium spp. <20 <10 0.11

Notes.a MGO (methylglyoxal) levels were analyzed against di-hydroxyacetone (DHA) and expressed as mg MGO per kg of honey.b Rate of production of H2O2 (hydrogen peroxide) is expressed as µmol/h in 1 mL of 10% w/v honey.* Information not available.

Other tested solutionsA series of other solutions were included for investigation alongside the honey samples:

(i) a sugar solution designed to mimic the concentration and composition of honey sugars

(45% glucose, 48% fructose, 1% sucrose) diluted as above for honey; (ii) MGO diluted in

TSB to concentrations similar to those present in the manuka-type honeys (100 mg/kg,

700 mg/kg and 900 mg/kg honey) to assess the effect of MGO alone on bacterial growth;

(iii) MGO diluted in sugar solution to the same concentration as (ii). MGO was obtained

as a ∼40% (w/w) solution in water (Sigma-Aldrich Co., MO, USA).

Hydrogen peroxide assayThe level of hydrogen peroxide produced by the NZ honey samples was determined using a

hydrogen peroxide/peroxidase assay kit (Amplex Red; Molecular Probes, Life Technologies

Corp., Carlsbad, CA, USA) as previously reported (Lu et al., 2013).

Bacterial strains and growth conditionsFour strains of S. aureus were examined. These include two laboratory reference strains:

NCTC 8325 (National Collection of Type Cultures) (Stepanovic et al., 2000) and ATCC

25923 (American Type Culture Collection) which are methicillin-sensitive, and two

clinical isolates: MW2 (Hospital-Acquired Methicillin-resistant Staphylococcus aureus,

HA-MRSA) (Baba et al., 2002) and USA300 (Community-Acquired Methicillin-resistant

Staphylococcus aureus, CA-MRSA) (Kazakova et al., 2005). All S. aureus strains were grown

in TSB at 37 ◦C. For optimal biofilm formation, 1% (w/v) glucose was added to this

medium (TSBG) except for strain NCTC 8325 which was found to produce optimal

biofilm in the absence of added glucose.

Susceptibility of S. aureus to NZ honeys: growth response assaysIn this study, growth response assays were carried out to assess whether the NZ honeys

affected cell growth of the different strains of S. aureus (at concentrations of 1–32%;

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 4/25

prepared as serial 2-fold dilutions in TSB(G)). Details of the growth assay methods are

described in our previous publication (Lu et al., 2013). TSB(G) media without honey

was included as a control. Unless otherwise stated, all assays were performed with three

biological replicates and three technical repeats of each replicate.

Biofilm formation assaysThe effects of NZ honeys and other solutions on S. aureus biofilm formation were

determined using crystal violet static biofilm formation assays in microtitre plates

according to published studies with the following modifications (Christensen et al.,

1985; Stepanovic et al., 2000). Crystal violet stains all biomass including live and dead

cells and the biofilm matrix. S. aureus strains were cultured in 2 mL of TSB(G) with

shaking (250 rpm) overnight at 37 ◦C . A suspension from the overnight culture was

then diluted to a cell density of approximately 107 CFU/mL in fresh TSB containing the

appropriate test solution (honey, sugar solution, MGO or MGO in combination with

sugar) to give a final volume of 150 µL. The suspension was added to each well of a 96-well

tissue-culture treated microtitre plate (BD Falcon, NJ, USA). Media-only and media with

the appropriate test solution without S. aureus inoculation were included as negative

controls. The microtitre plates were sealed with AeraSeal (Excel Scientific, CA, USA) and

incubated in a humidified incubator for 24 h at 37 ◦C. Following this, planktonic cell

growth was assessed by transferring the planktonic phase into a new 96-well microtitre

plate and reading the optical density at 595 nm with a microplate reader (VersaMax,

Molecular Devices, California, USA). This step was required as S. aureus forms biofilms

on the bottom of the microtitre plate wells, which interferes with optical density readings

of the planktonic culture. The microtitre plates with residual biofilm were then washed

three times with sterile phosphate buffered saline (PBS) to remove unattached cells and

air-dried at 65 ◦C for 1 h, to fix the S. aureus biofilm to the bottom of the well surface. The

plate was then stained with 0.2% (w/v) crystal violet at room temperature for 1 h, excess

crystal violet solution was decanted and the plates were washed as above with PBS. Stain

that was bound to the adherent biomass was resolubilized with 200 µL 33% acetic acid and

transferred into a new 96-well microtitre plate to measure the OD595.

Biofilm elimination assaysS. aureus biofilms were first formed in the wells of a 96-well microtitre plate for 24 h

at 37 ◦C as described above. Biofilms were then washed three times with PBS. Various

concentrations (0%–32% in two-fold serial dilutions) of honey and other test solutions

were then added to the established S. aureus biofilms. The assay plates were then incubated

for a further 24 h at 37 ◦C, and planktonic cell growth and biofilm mass were quantified as

described above.

Determination of bacterial cell viability in biofilmsCrystal violet stains all the components of the biofilm (Bauer et al., 2013). To quantify

the viability of cells within the S. aureus biofilms following honey treatment, we used a

BacTitre Glo Microbial Cell Viability Assay Kit (Promega, WI, USA), which measures

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ATP levels as a proxy for viability. The assay reagents lyse the bacterial cells to release

intracellular ATP, the levels of which are quantified via a luminescence-based luciferase

activity assay (Haddix et al., 2008; Junker & Clardy, 2007). The BacTitre Glo protocol

involved the same steps as crystal violet staining (above), however, instead of drying

and staining the biofilms, plates were incubated with BacTitre Glo reagent in TSB(G)

for 10 min at 37 ◦C in the dark. The contents of each well were then transferred into

white solid-bottom 96-well microtitre plates (Cellstar; Greiner Bio-one, France) for

luminescence measurement. Luminescence, which is proportional to the amount of ATP

produced by metabolically active cells, was recorded using a 96-well microplate reader

(Infinite 200Pro; TeCan, Mannedorf, Switzerland).

To ensure the validity of this assay, a standard curve was constructed to assess the

correlation between bacterial cell numbers and the luminescent signal in the biofilm.

This was performed on the untreated control (containing S. aureus in TSB(G) only).

Biofilms produced as above were washed and cells within the biofilm dispersed using a

small probe sonicator (Sonics and Materials VC-505) to enable quantification by direct

enumeration (Merritt, Kadouri & O’Toole, 2005). The recovered cell suspension was serially

diluted 10-fold and a 20 µL aliquot was plated on Tryptone Soya Agar (TSA) for CFU

determination. Luminescence of cells in the remaining suspension was assessed using

the BacTitre Glo kit. From this, a correlated standard curve was constructed between

calculated CFU/well and the relative luminescence readings. According to the standard

curve shown in Fig. 1, the detection limit of the BacTitre Glo is at a luminescence reading

below 1,000, which is equivalent to 103 CFU/well (linear range from 103–107 CFU/well).

An upper limit was not detected.

Visualizing live/dead stained S. aureus biofilms using confocallaser scanning microscope (CLSM)S. aureus biofilms were treated with TSB containing 1%, 2%, 16%, and 32% NZ honeys or

sugar solution for 24 h in black polystyrene 96-well microtitre plates with µClear bottoms

(Cellstar; Greiner Bio-One, France) as described above, except the biofilm mass was not

fixed by air-drying. The treated biofilm mass was washed three times with PBS and cells

within the biofilm structure were fluorescently stained with 2.5 µM Syto9 (Invitrogen, CA,

USA) and 4.3 µM propidium iodide (PI) (Becton Dickinson, NJ, USA), which identify

live and dead cells in the biofilm structure, respectively. After 30 min of incubation in the

dark at room temperature, the wells were washed thoroughly with PBS and fixed with 4%

paraformaldehyde (Sigma-Aldrich, MO, USA) for 15 min. The wells were then rinsed and

stored in PBS for imaging. Biofilms were imaged using confocal laser scanning microscopy

imaging (CLSM) on a Nikon A1 confocal microscope. The Syto9 and PI fluorophores

were excited at 488 nm and 561 nm, and the emissions were collected at 500–550 nm

and 570–620 nm, respectively. For quantitative analysis, at least eight separate CLSM

image stacks of each NZ honey treated biofilms were acquired with a resolution of 512 ×

512 pixels. Biofilm biomass was calculated using COMSTAT (Heydorn et al., 2000) and

is expressed as volume of the biofilm over the surface area (µm3/µm2). Representative

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Figure 1 Correlation of levels of intracellular ATP to colony forming units (CFU) in static biofilms ofS. aureus. Static biofilms of S. aureus were formed in the wells of a microtitre plate for 48 h (with mediareplenishment at 24 h). After removal of the biofilm from the wall of each well, intracellular ATP levelswere measured by the BacTitre Glo Viability Kit and CFU were determined for each well. The intracellularlevels of ATP are plotted as a function of CFU and validate that the BacTitre Glo Viability Kit can be usedas a surrogate measure of biofilm cell viability in subsequent assays.

presentation image stacks of each treatment were acquired at a resolution of 1024 ×

1024 pixels and three dimensional biofilm images reconstructed using NIS-elements

(Version10, Nikon Instruments Inc., USA). It should be noted that due to the incomplete

displacement of Syto9 by propidium iodide in dead cells that there will remain some Syto9

staining of dead cells. Therefore the absolute level of live cells detected in the Syto9 channel

will be somewhat overestimated using this co-staining method (Stocks, 2004).

Assaying honey resistance in cells recovered from biofilmsThe development of resistance is a great concern in clinical settings, where bacteria can

become resistant to inhibitory compounds after exposure to sub-inhibitory concentrations

(Cars & Odenholt-Tornqvist, 1993; Pankuch, Jacobs & Appelbaum, 1998). Planktonic cells

that appeared after 24 h manuka-type honey treatment of established biofilms were

assumed to have been released from the biofilm matrix. Cells recovered from biofilms

treated with sub-eliminatory concentrations of manuka-type honeys were collected and

tested for their ability to grow and form biofilms under the static growth conditions

described above. Cell growth and biofilm formation were defined as not detected when

the OD(x)–OD(media only blank) ≤ 0.1. Each experiment was performed with three biological

replicates and three technical repeats of each biological replicate.

Statistical analysisStatistical analysis to determine significant differences between treatments and among

honey samples were performed. First, data sets were checked for normality (Gaussian

Distribution) using the D’Agostino-Pearson normality test (alpha = 0.05) in GraphPad

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Table 2 Concentration of honey required to inhibit S. aureus growth.

Honeys NCTC 8325 ATCC 25923 MW2 USA300

Manuka honey 8* 8 8 8

Medihoney 8 8 8 8

Manuka/kanuka honey 16 16 16 16

Clover honey 32 32 32 32

Sugar solution >32 >32 32 >32

Notes.* All numbers in the table are honey concentrations (%, w/v).

Prism (versions 5 and 6). Once normality was confirmed, significant differences between

treatments and among honey samples were performed using One-Way ANOVA with Tukey

Test using the same software. Statistical significance was set at p < 0.05.

RESULTSThe effect of NZ manuka-type honeys on the planktonic growth ofS. aureusPlanktonic growth and biofilm mass were assessed to examine the ability of four NZ

honeys, three manuka-types and one clover, and a sugar solution to prevent biofilm

formation by different strains of S. aureus. Following 24 h incubation under static

conditions, S. aureus cells formed biofilms at the bottom of microtitre plates, with very

little or no planktonic growth detected, indicating that the concentration of honey

needed to prevent S. aureus planktonic cell growth could not be calculated under these

conditions. Shaking broth cultures were instead used to assess the effect of the treatments

on planktonic growth. The results are shown in Table 2. Planktonic growth of the four

S. aureus strains, NCTC 8325, ATCC 25923, MW2 and USA300, was completely inhibited

by 8% manuka honey and Medihoney, 16% manuka/kanuka honey and 32% clover honey.

The 32%, sugar solution was only effective at inhibiting growth of the MRSA strain MW2,

with no inhibition of growth of the other strains at the concentrations tested (1–32%).

These data are in agreement with the results of our previous study using the standard

S. aureus reference strain, ATCC 25923, which used a similar suite of honey types and the

same experimental conditions (Lu et al., 2013).

The effect of NZ manuka-type honeys on S. aureusbiofilm formationAll strains of S. aureus were assessed for their biofilm-forming ability after 24 h and 48 h.

Under static conditions, biofilm-forming ability varied between strains, with NCTC 8325

forming the most robust biofilms, and generating significantly more biofilm mass than

the other three tested strains (Figs. 2A and 2B). This was followed by ATCC 25923 and

USA300, with MW2 forming the thinnest biofilms (Figs. 2A and 2B; p < 0.05). The

effects of the four NZ honeys and the sugar solution on biofilm formation of strain NCTC

8325 are shown in Fig. 3A. Manuka honey was the most effective at preventing biofilm

formation by S. aureus NCTC 8325, resulting in ∼95% reduction (p < 0.001) in biofilm

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Figure 2 Quantification of biofilm formation by different strains of S. aureus. The ability of differentstrains of S. aureus to form biofilms on the plastic surface of tissue-culture treated 96-well microtitreplate was assessed in TSB(G) at 24 h and 48 h. Biofilm adherence was determined using a static biofilmformation assay over 24 h (A) and 48 h (with media replenished after 24 h incubation) (B). Biofilmformation was quantified by staining with 0.2% crystal violet solution and measured at an optical densityof 595 nm. Error bars represent ±standard deviation (SD) of three biological samples performed intriplicate, *** represents p < 0.001, compared to NCTC 8325, as assessed by One-Way ANOVA withTukey test after confirming normality of the data set for each treatment using the D’Agnostino-Pearsonnormality test.

formation at 8% (Fig. 3A) compared to the untreated (0%) control. At this concentration,

the other honeys and the sugar solution did not significantly reduce biofilm formation.

Medihoney and manuka/kanuka honey were highly effective at 16%, preventing biofilm

formation by ∼95% (p < 0.001). Clover honey was much less active and was less able to

prevent biofilm formation than the sugar solution, even at the highest concentration used

(32%).

For NCTC 8325, the addition of sub-inhibitory concentrations of manuka and

manuka/kanuka honey significantly enhanced biofilm formation, increasing it by 1.5-

and 2-fold, compared to the untreated control (p < 0.001). In contrast, Medihoney, clover

honey and the sugar solution did not enhance biofilm formation by strain NCTC 8325 at

any concentration tested.

S. aureus strain ATCC 25923, which is a standard clinical reference strain, produced

similar results to NCTC 8325, including the enhancement of biofilm formation following

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Figure 3 Effects of NZ honeys and sugar on S. aureus biofilm formation. S. aureus biofilms wereallowed to form in the presence of four different NZ honey types (manuka, Medihoney, manuka/kanukaor clover) or a sugar solution. Biofilm formation was assessed using a static biofilm formation assaywith crystal violet staining to quantify biomass. S. aureus strains are: (A) NCTC 8325; (B) ATCC 25923;(C) MW2 (HA-MRSA) and (D) USA300 (CA-MRSA). Biofilm formation is expressed as a percentagerelative to that produced by the untreated control, which is set at 100%. Error bars represent ±standarddeviation (SD) of three biological samples performed in triplicate. Statistical significance (p < 0.05)was assessed by One-Way ANOVA with Tukey test after confirming normality of the data set for eachtreatment using the D’Agnostino-Pearson normality test.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 10/25

sub-inhibitory honey treatment (Fig. 3B). The hospital-acquired MRSA strain MW2—the

weakest biofilm former out of all four tested strains (Figs. 2A and 2B; p < 0.05)—displayed

a very sensitive profile to all of the NZ honeys and the sugar solution at all tested

concentrations (Fig. 3C). Even with only 1% honey or sugar solution, a ∼50% reduction in

biofilm formation was observed for MW2 (p < 0.001). At higher concentrations (≥8%),

all four NZ honeys were significantly more effective than the sugar solution at preventing

MW2 biofilms. The other MRSA strain, USA 300, responded similarly to NCTC 8325, with

approximately the same concentrations of manuka-type honey being required to reduce

biofilm formation by ∼95%. However, unlike NCTC 8325, sub-inhibitory concentrations

of manuka-type honey reduced biofilm formation of USA 300 rather than enhancing

it (e.g., 4% manuka-type honeys exhibited ∼50–80% biofilm inhibition of USA 300).

Moreover, in USA300, biofilm formation was not affected by the sugar solution at any

tested concentration.

The results above can be summarize as follows: (i) all three manuka-type honeys are

effective at inhibiting biofilm formation of a range of of MSSA and MRSA strains; with

(monofloral) manuka honey being generally more effective than the other maunka-type

honeys; and (ii) the manuka-type honeys are generally more effective than clover honey

and the isotonic sugar solution, although clover honey was just as inhibitory as the

manuka-type honeys for the weakest biofilm former, S. aureus MW2.

The effect of MGO on S. aureus biofilm preventionMGO is a principle antibacterial component of manuka honey responsible for its

inhibitory effects on the growth of S. aureus and other bacterial species. This is evidenced

by the correlation between the MGO level and the proportion of manuka-derived honey

in a honey blend (Jervis-Bardy et al., 2011; Lu et al., 2013). To determine whether MGO is

solely responsible for the inhibitory effect of the three manuka-type honeys on S. aureus

biofilm formation, biofilm assays were performed using MGO at equivalent concentrations

to those present in each of the manuka-based honey samples, with and without the

addition of the sugar solution (Fig. 4). S. aureus NCTC 8325 biofilm formation was not

significantly (p > 0.05) affected by MGO at concentrations equivalent to that present in

1–16% manuka-type honeys. MGO at medium (700 mg/kg) and high (900 mg/kg) levels

at the equivalent concentration to 32% manuka-kanuka honey and Medihoney prevented

approximately 50% and 75% biofilm formation, respectively (p < 0.05). The addition of

the sugar solution to MGO at the same levels present in 16% of all three manuka-type

honeys, led to a dramatic decrease (∼95%) in biofilm formation.

The effect of NZ manuka-type honeys on established S. aureusbiofilmsBacterial biofilms are usually already established in open, chronic wounds prior to

presentation to the clinic for medical treatment. We therefore assessed the ability of the

four NZ honeys to remove established biofilms produced by the four strains of S. aureus.

These results are presented in Fig. 5, with coloured lines showing biofilm mass present

following treatment with different concentrations of the various honey types. While

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Figure 4 Effects of MGO on S. aureus biofilm formation. Biofilm formation by S. aureus NCTC 8325grown in the presence of MGO and MGO plus sugar solution. MGO stock solutions were prepared tocorrespond to the MGO levels in undiluted manuka-type honeys (100 mg/kg of manuka/kanuka honey,700 mg/kg of Medihoney, and 900 mg/kg of manuka-honey; Table 1). Biofilm formation was assessedusing the described static assay with crystal violet staining to quantify biomass. Biofilm formation isexpressed as a percentage relative to the untreated control, which is set at 100%. Error bars represent±standard deviation (SD) of three biological samples performed in triplicate. Statistical significance(p < 0.05) was assessed by One-Way ANOVA with Tukey test after confirming normality of the dataset for each treatment using the D’Agnostino-Pearson normality test.

there was variation among the S. aureus strains in their response to the different honeys,

there are some important general trends. First, manuka honey was consistently the most

effective at removing biofilm, eliminating almost all of the established S. aureus biofilms

at concentrations of 16%–32%, (p < 0.001 compared to the untreated control sample;

Fig. 5 top panel, orange lines). Second, Medihoney and manuka/kanuka honey were also

effective at these concentrations for some S. aureus strains, but only consistently effective

across all four strains at 32% (Fig. 5, blue and green lines). Third, both the clover honey

and the sugar solution did not significantly reduce (p > 0.5) established biofilm mass until

their concentration reached 32%. However, the sugar solution did not remove the USA 300

biofilm, with no significant reduction in biofilm mass at 32% (Fig. 5, purple line).

NCTC 8325, the most efficient biofilm former out of all tested strains, gave a slightly

different response toward honey treatment compared to the other three strains. Significant

biofilm enhancement occurred in this strain at sub-inhibitory concentrations of manuka

honey (1–2%) and manuka/kanuka honey (1–4%) (p < 0.001; Fig. 5). In addition, this

strain was the least sensitive to the manuka-type honeys. For example, at 8% manuka

honey treatment, the NCTC 8325 biofilm mass remained similar to the untreated control

(p > 0.05; Fig. 5), while the biofilms produced by the other three strains were significantly

reduced at this concentration (p < 0.001; Fig. 5).

The effect of NZ manuka-type honeys on cell viability withinS. aureus biofilmsElimination of biofilm mass was assessed using crystal violet, a cationic dye that stains

all the components of the biofilm. However, this assay cannot assess the viability of cells

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 12/25

Figure 5 Effects of NZ honeys on established S. aureus biofilms and cell viability within the biofilms. Established S. aureus NCTC 8325, ATCC25923, MW2 and USA300 biofilms were treated with NZ honeys – manuka, Medihoney, manuka/kanuka, clover, and a sugar solution. The remainingbiofilm masses were quantified using crystal violet staining (left y-axis) and cell viability within these remaining biofilms were assessed using theBacTitre Glo Viability Kit (right y-axis). Error bars represent ±standard deviation (SD) of three biological samples performed in triplicate Statisticalsignificance (p < 0.05) was assessed by One-Way ANOVA with Tukey test after confirming normality of the data set for each treatment using theD’Agnostino-Pearson normality test.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 13/25

remaining within the biofilm structure (Bauer et al., 2013). To determine this, we used a

BacTitre Glo assay, which measures ATP levels as a proxy for viability. Side-by-side CFU

measurements showed that the level of ATP detected in these assays was proportional to the

count of viable cells per well (ranging from 103 to 107 CFU/well) (Fig. 1).

The viability of cells remaining in the biofilm after the various treatments is shown

in Fig. 5. In general, cell viability decreased in proportion to the elimination of biofilm

biomass (Fig. 5, black lines). However, several exceptions to this general trend were

observed. In some cases, biofilm biomass increased but cell viability did not, e.g., NCTC

8325 biofilms with low concentrations of manuka (2%) and manuka/kanuka honey

(1–4%) (Fig. 5). In others, biofilm biomass remained relatively constant while cell viability

increased, e.g., NCTC 8325 with 1–4% Medihoney (p < 0.05; Fig. 5), and ATCC 25923

with 4% and 8% clover honey. Another deviation from the general trend was a significant

reduction of cell viability while biofilm biomass remained unaffected, seen for NCTC 8325

and USA 300 with 4% and 8% manuka honey treatment (Fig. 5; p < 0.05). This emphasizes

the importance of assessing viability alongside crystal violet assays for biofilm assessment.

Overall, at concentrations easily attainable in the clinic, the tested four NZ honeys

were effective at eliminating biofilm biomass and at killing both MSSA and MRSA

S. aureus cells in the residual biofilm. Among the honey types, manuka honey was the

most effective, where the elimination of biofilm biomass largely paralleled the reduction

in viability. Following treatment with 8% manuka honey only ∼10% of cells were viable

in the remaining ATCC 25923 and USA 300 biofilms, compared to the untreated control

(i.e., 0% honey), and no generation of ATP could be detected from MW2 (Fig. 5). This

is similar to the degree of biofilm biomass removal, where 85–98% of biofilm biomass

was removed following 8% manuka honey treatment. Although NCTC 8325 biofilm

biomass was seemingly unaffected at 8% manuka honey compared to the untreated control

(Fig. 5), the number of viable cells detected within this biofilm was drastically reduced by

approximately 80% (p < 0.001; Fig. 5).

The effect of MGO on established S. aureus biofilmsTo assess the contribution of MGO alone, as well as MGO plus sugar, to biofilm removal,

these components were tested on established S. aureus NCTC 8325 biofilms (Fig. 6). MGO

levels equivalent to the presence of 1–8% manuka/kanuka honey (Table 1) caused biofilm

biomass to increase approximately 2-fold, relative to the untreated control (p < 0.001).

However, the established biofilm biomass was not reduced significantly (p > 0.05), for any

of the tested concentrations (1–32%) of MGO by itself, or in combination with the sugar

solution. Thus, neither MGO nor the combination of MGO with sugar is solely responsible

for the elimination of biofilms observed with these manuka-type honeys.

Visualizing the effects of NZ manuka-type honeys on establishedS. aureus biofilmsTo assess the effect of the NZ honeys on S. aureus NCTC 8325 biofilms at the cellular level,

we used confocal laser scanning microscopy (CLSM) of biofilms stained with fluorescent

dyes for the detection of live and dead bacteria. This allows both the visualization of

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 14/25

Figure 6 Effects of MGO on established S. aureus biofilms. S. aureus NCTC 8325 biofilms were treatedwith MGO and a combination of MGO and the sugar solution. MGO stock solutions were prepared tocorrespond to the MGO levels in undiluted honey (100 mg/kg of manuka/kanuka honey, 700 mg/kg ofMedihoney, and 900 mg/kg of manuka honey; Table 1). The crystal violet stained residual biofilm massafter 24 h treatment was quantified using optical density (OD595 nm). Error bars represent ±standarddeviation (SD) of three biological samples performed in triplicate. Statistical significance (p < 0.05)was assessed by One-Way ANOVA with Tukey test after confirming normality of the data set for eachtreatment using the D’Agnostino-Pearson normality test.

individual cells within the biofilm in three dimensions and the effect of treatments on cell

viability to be determined. Treatment by sub-inhibitory (1% and 2%) and inhibitory (16%

and 32%) concentrations of NZ honeys was visualized by viewing fluorescently-labelled

live (Syto9; green) and dead (propidium iodide; red) cells. Representative images of each

treatment are presented in Fig. 7 and quantification of live and dead cell biofilm biomass

for several samples for each treatment is shown in Fig. 8. In general, the established

biofilm biomass decreased with increasing concentrations of manuka-type honey. More

specifically, manuka honeys were effective in reducing the live cells in established S. aureus

biofilms. Sub-inhibitory concentrations of all the manuka-type honeys (1% and 2%) and

the sugar solution did not reduce the amount of biomass compared to the non-treated

control cells (Fig. 8). This is shown in Fig. 7 where the untreated control cells displayed a

green (live-cell) lawn that covered nearly the entire surface and this remained following

treatment with 1% and 2% manuka-type honeys. At concentrations of 16% and 32%, the

manuka-type honeys substantially reduced the density and depth of the biofilm, along with

the amount of live cells, compared to the untreated control (Figs. 7 and 8). For example, the

32% manuka honey significantly reduced the Syto9 stained (live) biofilm biomass to 10%

(p < 0.001) compared to the non-treated live biofilm biomass (Fig. 8).

Only small micro-colonies were present following treatment with 32% manuka honey,

and the colour of the biofilms was predominantly yellow (where both the green and red dye

were retained within cells), indicating mostly dead cells. In contrast, 32% clover honey and

sugar solution reduced the total biomass by a maximum of 30% (p < 0.001) compared to

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 15/25

Figure 7 Live/dead staining of different honey treated established biofilms. Established biofilmsproduced by S. aureus NCTC 8325 were treated with TSB containing honey (manuka, Medihoney,manuka/kanuka or clover) or sugar solution at 1%, 2%, 16%, and 32% (w/vol). Syto9 (green; viablecells) and propidium iodine (red; dead cells) stained images were acquired using Nikon A1 ConfocalLaser Scanning Microscope. The 3D- images were reconstructed using NIS-elements (version 10). Scalebar represents 50 µm.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 16/25

Figure 8 Quantitative analysis of live/dead stained honey treated biofilms. The established S. aureusNCTC 8325 biofilm was treated with New Zealand honeys (manuka honey, Medihoney, manuka/kanukahoney, and clover honey) and a sugar solution at 1%, 2%, 16%, and 32% (w/v) concentrations. Biofilmswere co-stained with Syto9 (S, viable cells) and propidium iodide (P, dead cells) and analyzed usingCOMSTAT. The estimated live (S) and dead (P) biomass (volume of the biofilm over the surface area(µm3/µm2)) are expressed as a percentage of the non-treated control live and dead biomass, which is setat 100%. Error bars represent ±standard deviation (SD) of three biological samples where eight repre-sentative images were acquired. Statistical significance (p < 0.05) was assessed by One-Way ANOVA withTukey test after confirming normality of the data set for each treatment using the D’Agnostino-Pearsonnormality test.

the non-treated control (Fig. 8). This result corresponds to the 3D reconstructed images,

where the Syto9 stained cells remained dominant after treatment (Fig. 7). At 32%, clover

honey or sugar solution, a substantially larger Syto9 stained (live) lawn remained in

comparison to the 32% manuka-type honeys, although the biomass was less confluent

than in the untreated control. These results are consistent with the results obtained with

the crystal violet stained biofilm biomass and ATP viability assays.

Assessing the development of resistance to manuka-type honeysin S. aureus biofilmsBacteria that are exposed to sub-inhibitory concentrations of antimicrobial agents

generally develop resistance to these agents (Braoudaki & Hilton, 2004; Davies, Spiegelman

& Yim, 2006). The ability of cells released from S. aureus NCTC 8325 biofilms to develop

resistance after exposure to sub-inhibitory concentrations of honey was investigated, and

the results are summarized in Table 3. All cells recovered from the S. aureus biofilm after

24 h with 8% of all three manuka-type honeys were viable and able to form biofilms

in media (TSB). However, they were unable to grow planktonically when subsequently

exposed to 8% manuka, or 16% Medihoney and manuka/kanuka honey (the MIC levels

for these honeys). Biofilm formation was also inhibited by 8% manuka honey, and by

16% Medihoney and manuka/kanuka honey. These growth- and biofilm-inhibitory

concentrations of manuka-type honeys are the same as those observed for cells that had

not previously been treated with these honeys. These results indicating that planktonic cells

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 17/25

Table 3 Resistance of S. aureus cells recovered from biofilms after 8% manuka honey treatmentsa.

Honey (%) Type of assay Manuka honey Medihoney Manuka/kanukahoney

Growth√ √ √

0Biofilm formation

√ √ √

Growth × ×√

8Biofilm formation ×

√ √

Growth × × ×16

Biofilm formation × × ×

Notes.a A tick means that there was normal growth or biofilm formation and a cross means that there was no growth or no

bioflm formation.

released from the biofilms with exposure to sub-inhibitory concentrations did not acquire

resistance to the same honey treatment.

DISCUSSIONChronic wounds are costly and difficult to treat (Hoyle & Costerton, 1991; Ranall et al.,

2012; Sen et al., 2009), and bacterial biofilms are important contributors to the delay in

healing. Honey is a promising alternative treatment for these wounds, and studies have

indicated that it is able to prevent bacterial biofilms and eliminate established biofilms

in vitro (Alandejani et al., 2008; Maddocks et al., 2013; Maddocks et al., 2012; Majtan

et al., 2013). However, the effective concentration of honey reported by these studies

varies significantly, making it hard to establish a foundation for the efficacy of honey on

chronic wound-associated bacterial biofilms in the clinic. This is probably largely due to

the fact that, in most of these studies, very little information is reported on the honey itself,

including the floral source, geographic location, storage conditions, and the level of the

two principle antibacterial components, MGO and hydrogen peroxide. Here we utilize a

suite of well-defined NZ honeys, including manuka-type honeys (manuka, Medihoney

and manuka/kanuka honey) and clover honey, to investigate their anti-biofilm activity

on a range of S. aureus biofilms that differ in their ability to form biofilms. We show

that manuka-type honeys can be used to kill all MSSA and MRSA cells when present as

a biofilm in a chronic wound, supporting the use of this honey as an effective topical

treatment for chronic wound infections.

Our study has shown that prevention of S. aureus biofilm formation occurred at honey

concentrations that also inhibit planktonic growth (Figs. 3A–3D; Table 2), suggesting that

biofilm prevention was a consequence of planktonic growth inhibition, as opposed to any

specific effects on biofilm development. Other studies have also shown that manuka-type

honeys can inhibit bacterial biofilm formation, however, the concentrations required were

higher than those reported to inhibit growth (Alandejani et al., 2008; Maddocks et al.,

2013; Maddocks et al., 2012; Majtan et al., 2013).

We found that higher concentrations of all honeys were necessary to eliminate

established biofilms compared to those needed for prevention, as assessed by both

quantification of biofilm biomass and cell viability. Manuka honey was the most effective,

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 18/25

closely followed by both Medihoney and manuka/kanuka honey. Elimination of biofilms

was visually confirmed using CLSM of fluorescently-stained live and dead cells. The sugar

content of honey clearly mediates some effect, as sugar solution and clover honey were able

to eliminate established biofilms at high concentrations (32%), as has been shown in other

studies (Chirife et al., 1983; Chirife, Scarmato & Herszage, 1982). However, manuka-type

honeys consistently achieved biofilm elimination at lower concentrations, suggesting

that components specifically within manuka-type honeys contribute towards biofilm

elimination. The concentrations of manuka-type honeys that show significant anti-biofilm

activity are easily achievable in the clinic, since honey dressings typically contain >80%

honey (Cooper et al., 2010).

The use of assays for total biofilm biomass and cell viability to examine the effects of the

various treatments on biofilm elimination afforded some other interesting observations.

We observed that in some cases, sub-inhibitory concentrations of two of the manuka-type

honeys enhanced biofilm formation; however, cell viability did not increase. This could

be due to a stress response, as has been previously observed when bacteria are exposed to

sub-inhibitory concentrations of antibiotics (Haddadin et al., 2010; Kaplan et al., 2012;

Mirani & Jamil, 2011; Subrt, Mesak & Davies, 2011). In other cases, no reduction of biofilm

biomass was observed but cell viability was significantly reduced. This suggests that unlike

antibiotics, the manuka-type honeys (or active components therein) are able to penetrate

through the biofilm matrix, killing the bacterial cells whilst leaving intact matrix.

It is believed that MGO is the primary component in manuka-type honeys responsible

for its anti-biofilm activity (Jervis-Bardy et al., 2011; Kilty et al., 2011). The effectiveness of

the different manuka-type honeys tested here did increase with MGO content. However,

the same degree of biofilm prevention and elimination could not be reproduced with

equivalent amounts of MGO either alone or in combination with sugar. In the case of

prevention, MGO alone was generally ineffective, although a significant amount of biofilm

prevention was achieved in combination with sugar. This suggests that the MGO and sugar

do contribute to biofilm prevention, but their effects are not as strong as those observed

with manuka honey.

Unlike the three NZ manuka-type honeys, neither MGO alone nor MGO with sugar at

honey-equivalent concentrations showed significant S. aureus biofilm elimination. This

indicates that the ability of manuka-type honeys to eliminate biofilms of this organism

is due to one or more components present in the honey other than MGO and sugar,

such as low pH, hydrogen peroxide, phenolics and other unknown components (Jagani,

Chelikani & Kim, 2009; Jervis-Bardy et al., 2011; Kilty et al., 2011; Zmantar et al., 2010).

Interestingly, while the kanuka/manuka honey had a relatively high rate of hydrogen

peroxide production compared to the manuka and Medihoney (Table 1), but low MGO

levels, it was not any more active against biofilms of S. aureus. This suggests that, at least

for this organism, hydrogen peroxide within these manuka-type honeys does not provide

significant anti-biofilm activity.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 19/25

CONCLUSIONSThis study is the first to use a suite of well-characterized manuka-type honeys against a

range of strains of S. aureus that differ in their ability to form biofilms. We demonstrate

that: (1) at very low levels, some honeys can enhance biofilm formation, presumably

by evoking a stress response similar to that seen with some antibiotics; (2) the ability

to prevent or eliminate biofilms is influenced by MGO levels and the presence of sugar,

but these alone do not account for all of the anti-biofilm effect; (3) honey is able to

reduce biofilm mass and also to kill cells that remain embedded in the biofilm matrix;

and (4) planktonic cells released from biofilms following honey treatment do not have

elevated resistance to honey. Taken together our results show that if used at an appropriate

therapeutic level, manuka-type honey can be used to kill S. aureus when present as a

biofilm in a chronic wound, supporting the use of this honey as an effective topical

treatment for chronic wound infections.

ACKNOWLEDGEMENTSCSLM was performed at the UTS Microbial Imaging Facility.

ADDITIONAL INFORMATION AND DECLARATIONS

FundingFunding sources include: Comvita NZ Ltd, The Australian Research Council (ARC Linkage

Project LP0990949), CBW was supported by an Australian National Health and Medical

Research Council Senior Research Fellowship (571905). LT was supported by a University

of Technology, Sydney Chancellor’s Postdoctoral Fellowship. The funders had no role

in study design, data collection and analysis, decision to publish, or preparation of the

manuscript.

Grant DisclosuresThe following grant information was disclosed by the authors:

Australian Research Council Linkage Project grant: LP0990949.

Australian National Health and Medical Research Council Senior Research Fellowship:

571905.

Competing InterestsRalf C. Schlothauer is an employee of Comvita New Zealand (NZ) Limited which trades

in medical grade manuka honey (Medihoney). Comvita NZ Ltd. have partially funded

the work through a contribution to Linkage Project LP0990949 funded by the Australian

Research Council. Chief Investigators on this project include Elizabeth J. Harry, Cynthia B.

Whitchurch, Lynne Turnbull, Dee A. Carter and Partner Investigator Ralf C. Schlothauer.

Our competing interests do not alter our adherence to all the PeerJ policies on sharing

data and materials. We also note that co-author Associate Professor Dee A. Carter is an

Academic Editor for PeerJ.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 20/25

Author Contributions• Jing Lu performed the experiments, analyzed the data, wrote the paper, prepared figures

and tables, reviewed drafts of the paper.

• Lynne Turnbull and Cynthia B. Whitchurch conceived and designed the experiments,

analyzed the data, contributed reagents/materials/analysis tools, reviewed drafts of the

paper, revised the manuscript.

• Catherine M. Burke analyzed the data, wrote the paper, reviewed drafts of the paper.

• Michael Liu analyzed the data, wrote the paper, reviewed drafts of the paper.

• Dee A. Carter analyzed the data, contributed reagents/materials/analysis tools, wrote the

paper, reviewed drafts of the paper.

• Ralf C. Schlothauer conceived and designed the experiments, analyzed the data,

contributed reagents/materials/analysis tools, revised the manuscript.

• Elizabeth J. Harry conceived and designed the experiments, analyzed the data,

contributed reagents/materials/analysis tools, wrote the paper, reviewed drafts of the

paper, revised the manuscript and coordinated the research.

REFERENCESAdams CJ, Boult CH, Deadman BJ, Farr JM, Grainger MNC, Manley-Harris M, Snow MJ.

2008. Isolation by HPLC and characterisation of the bioactive fraction of NewZealand manuka (Leptospermum scoparium) honey. Carbohydrate Research 343:651–659DOI 10.1016/j.carres.2007.12.011.

Al-Waili N, Al Ghamdi A, Ansari MJ, Al-Attar Y, Osman A, Salom K. 2013. Differences incomposition of honey samples and their impact on the antimicrobial activities againstdrug multiresistant bacteria and pathogenic fungi. Archives of Medical Research 44:307–316DOI 10.1016/j.arcmed.2013.04.009.

Al-Waili N, Salom K, Al-Ghamdi AA. 2011. Honey for wound healing, ulcers, and burns;data supporting its use in clinical practice. The Scientific World Journal 11:766–787DOI 10.1100/tsw.2011.78.

Alandejani T, Marsan JG, Ferris W, Slinger R, Chan F. 2008. Effectiveness of honey onS. aureus and P. aeruginosa biofilms. Otolaryngology – Head and Neck Surgery 139:P107–P107DOI 10.1016/j.otohns.2008.05.541.

Allen KL, Molan PC, Reid GM. 1991. A survey of the antibacterial activity of some New Zealandhoneys. Journal of Pharmacy and Pharmacology 43:817–822DOI 10.1111/j.2042-7158.1991.tb03186.x.

Baba T, Takeuchi F, Kuroda M, Yuzawa H, Aoki K, Oguchi A, Nagai Y, Iwama N, Asano K,Naimi T, Kuroda H, Cui L, Yamamoto K, Hiramatsu K. 2002. Genome and virulencedeterminants of high virulence community-acquired MRSA. Lancet 359:1819–1827DOI 10.1016/S0140-6736(02)08713-5.

Battin TJ, Sloan WT, Kjelleberg S, Daims H, Head IM, Curtis TP, Eberl L. 2007. Microbiallandscapes: new paths to biofilm research. Nature Reviews Microbiology 5:76–81DOI 10.1038/nrmicro1556.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 21/25

Bauer J, Siala W, Tulkens PM, Van Bambeke F. 2013. A combined pharmacodynamic quantitativeand qualitative model reveals the potent activity of Daptomycin and Delafloxacin againstStaphylococcus aureus biofilms. Antimicrobial Agents and Chemotherapy 57:2726–2737DOI 10.1128/AAC.00181-13.

Blair SE, Cokcetin NN, Harry EJ, Carter DA. 2009. The unusual antibacterial activity ofmedical-grade Leptospermum honey: antibacterial spectrum, resistance and transcriptomeanalysis. European Journal of Clinical Microbiology and Infectious Diseases 28:1199–1208DOI 10.1007/s10096-009-0763-z.

Braoudaki M, Hilton AC. 2004. Adaptive resistance to biocides in Salmonella enterica andEscherichia coli O157 and cross-resistance to antimicrobial agents. Journal of ClinicalMicrobiology 42:73–78 DOI 10.1128/JCM.42.1.73-78.2004.

Cars O, Odenholt-Tornqvist I. 1993. The post-antibiotic sub-MIC effect in vitro and in vivo.Journal of Antimicrobial Chemotherapy 31(Suppl D):159–166 DOI 10.1093/jac/31.suppl D.159.

Chirife J, Herszage L, Joseph A, Kohn ES. 1983. In vitro study of bacterial growth inhibition inconcentrated sugar solutions: microbiological basis for the use of sugar in treating infectedwounds. Antimicrobial Agents and Chemotherapy 23:766–773 DOI 10.1128/AAC.23.5.766.

Chirife J, Scarmato G, Herszage L. 1982. Scientific basis for use of granulated sugar in treatmentof infected wounds. Lancet 1:560–561 DOI 10.1016/S0140-6736(82)92065-7.

Christensen GD, Simpson WA, Younger JJ, Baddour LM, Barrett FF, Melton DM, Beachey EH.1985. Adherence of coagulase-negative staphylococci to plastic tissue culture plates: aquantitative model for the adherence of staphylococci to medical devices. Journal of ClinicalMicrobiology 22:996–1006.

Colsky AS, Kirsner RS, Kerdel FA. 1998. Analysis of antibiotic susceptibilities of skin wound florain hospitalized dermatology patients. The crisis of antibiotic resistance has come to the surface.Archives of Dermatology 134:1006–1009 DOI 10.1001/archderm.134.8.1006.

Cooper R. 2008. Using honey to inhibit wound pathogens. Nursing Times 104(46):48–49.

Cooper RA, Jenkins L, Henriques AF, Duggan RS, Burton NF. 2010. Absence of bacterialresistance to medical-grade manuka honey. European Journal of Clinical Microbiology andInfectious Diseases 29:1237–1241 DOI 10.1007/s10096-010-0992-1.

Costerton W, Veeh R, Shirtliff M, Pasmore M, Post C, Ehrlich G. 2003. The application of biofilmscience to the study and control of chronic bacterial infections. Journal of Clinical Investigation112:1466–1477 DOI 10.1172/JCI200320365.

Davies J, Spiegelman GB, Yim G. 2006. The world of subinhibitory antibiotic concentrations.Current Opinion in Microbiology 9:445–453 DOI 10.1016/j.mib.2006.08.006.

Engemann JJ, Carmeli Y, Cosgrove SE, Fowler VG, Bronstein MZ, Trivette SL, Briggs JP,Sexton DJ, Kaye KS. 2003. Adverse clinical and economic outcomes attributable to methicillinresistance among patients with Staphylococcus aureus surgical site infection. Clinical InfectiousDiseases 36:592–598 DOI 10.1086/367653.

Forrest RD. 1982. Early history of wound treatment. Journal of the Royal Society of Medicine75:198–205.

Fux CA, Costerton JW, Stewart PS, Stoodley P. 2005. Survival strategies of infectious biofilms.Trends in Microbiology 13:34–40 DOI 10.1016/j.tim.2004.11.010.

Gresley AL, Kenny J, Cassar C, Kelly A, Sinclair A, Fielder MD. 2012. The application of highresolution diffusion NMR to the analysis of manuka honey. Food Chemistry 135:2879–2886DOI 10.1016/j.foodchem.2012.07.072.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 22/25

Haddadin RN, Saleh S, Al-Adham IS, Buultjens TE, Collier PJ. 2010. The effect of subminimalinhibitory concentrations of antibiotics on virulence factors expressed by Staphylococcus aureusbiofilms. Journal of Applied Microbiology 108:1281–1291DOI 10.1111/j.1365-2672.2009.04529.x.

Haddix PL, Jones S, Patel P, Burnham S, Knights K, Powell JN, LaForm A. 2008. Kinetic analysisof growth rate, ATP, and pigmentation suggests an energy-spilling function for the pigmentprodigiosin of Serratia marcescens. Journal of Bacteriology 190:7453–7463DOI 10.1128/JB.00909-08.

Heydorn A, Nielsen AT, Hentzer M, Sternberg C, Givskov M, Ersboll BK, Molin S. 2000.Quantification of biofilm structures by the novel computer program COMSTAT. Microbiology146(Pt 10):2395–2407.

Hoyle BD, Costerton JW. 1991. Bacterial resistance to antibiotics: the role of biofilms. Progress inDrug Research 37:91–105.

Jagani S, Chelikani R, Kim DS. 2009. Effects of phenol and natural phenolic compounds onbiofilm formation by Pseudomonas aeruginosa. Biofouling 25:321–324DOI 10.1080/08927010802660854.

Jervis-Bardy J, Foreman A, Bray S, Tan L, Wormald PJ. 2011. Methylglyoxal-infused honeymimics the anti-Staphylococcus aureus biofilm activity of manuka honey: potential implicationin chronic rhinosinusitis. Laryngoscope 121:1104–1107 DOI 10.1002/lary.21717.

Junker LM, Clardy J. 2007. High-throughput screens for small-molecule inhibitors ofPseudomonas aeruginosa biofilm development. Antimicrobial Agents and Chemotherapy51:3582–3590 DOI 10.1128/AAC.00506-07.

Kaplan JB, Izano EA, Gopal P, Karwacki MT, Kim S, Bose JL, Bayles KW, Horswill AR. 2012.Low levels of β-lactam antibiotics induce extracellular DNA release and biofilm formation inStaphylococcus aureus. mBio 3:e00198-12 DOI 10.1128/mBio.00198-12.

Kazakova SV, Hageman JC, Matava M, Srinivasan A, Phelan L, Garfinkel B, Boo T, McAllister S,Anderson J, Jensen B, Dodson D, Lonsway D, McDougal LK, Arduino M, Fraser VJ,Killgore G, Tenover FC, Cody S, Jernigan DB. 2005. A clone of methicillin-resistantStaphylococcus aureus among professional football players. New England Journal of Medicine352:468–475 DOI 10.1056/NEJMoa042859.

Kilty SJ, Duval M, Chan FT, Ferris W, Slinger R. 2011. Methylglyoxal: (active agent of manukahoney) in vitro activity against bacterial biofilms. International Forum of Allergy & Rhinology1:348–350 DOI 10.1002/alr.20073.

Kwakman PH, te Velde AA, de Boer L, Speijer D, Vandenbroucke-Grauls CM, Zaat SA. 2010.How honey kills bacteria. FASEB Journal 24:2576–2582 DOI 10.1096/fj.09-150789.

Kwakman PH, Te Velde AA, de Boer L, Vandenbroucke-Grauls CM, Zaat SA. 2011. Two majormedicinal honeys have different mechanisms of bactericidal activity. PLoS ONE 6:e17709DOI 10.1371/journal.pone.0017709.

Kwakman PH, Zaat SA. 2012. Antibacterial components of honey. IUBMB Life 64:48–55DOI 10.1002/iub.578.

Lu J, Carter DA, Turnbull L, Rosendale D, Hedderley D, Stephens J, Gannabathula S,Steinhorn G, Schlothauer RC, Whitchurch CB, Harry EJ. 2013. The effect of New Zealandkanuka, manuka and clover honeys on bacterial growth dynamics and cellular morphologyvaries according to the species. PLoS ONE 8:e55898 DOI 10.1371/journal.pone.0055898.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 23/25

Maddocks SE, Jenkins RE, Rowlands RS, Purdy KJ, Cooper RA. 2013. Manuka honey inhibitsadhesion and invasion of medically important wound bacteria in vitro. Future Microbiology8:1523–1536 DOI 10.2217/fmb.13.126.

Maddocks SE, Lopez MS, Rowlands RS, Cooper RA. 2012. Manuka honey inhibits thedevelopment of Streptococcus pyogenes biofilms and causes reduced expression of twofibronectin binding proteins. Microbiology 158:781–790 DOI 10.1099/mic.0.053959-0.

Majtan J, Bohova J, Horniackova M, Klaudiny J, Majtan V. 2013. Anti-biofilm effects of honeyagainst wound pathogens proteus mirabilis and enterobacter cloacae. Phytotherapy Research28:69–75 DOI 10.1002/ptr.4957.

Mavric E, Wittmann S, Barth G, Henle T. 2008. Identification and quantification of methylglyoxalas the dominant antibacterial constituent of Manuka (Leptospermum scoparium) honeys fromNew Zealand. Molecular Nutrition & Food Research 52:483–489 DOI 10.1002/mnfr.200700282.

McCarty SM, Cochrane CA, Clegg PD, Percival SL. 2012. The role of endogenous and exogenousenzymes in chronic wounds: a focus on the implications of aberrant levels of both hostand bacterial proteases in wound healing. Wound Repair and Regeneration 20:125–136DOI 10.1111/j.1524-475X.2012.00763.x.

Merritt JH, Kadouri DE, O’Toole GA. 2005. Growing and analyzing static biofilms.Current Protocols in Microbiology 22:1B.1.1–1B.1.18. Chapter 1:Unit 1B.1DOI 10.1002/9780471729259.mc01b01s00.

Mirani ZA, Jamil N. 2011. Effect of sub-lethal doses of vancomycin and oxacillin on biofilmformation by vancomycin intermediate resistant Staphylococcus aureus. Journal of BasicMicrobiology 51:191–195 DOI 10.1002/jobm.201000221.

Molan PC. 1999. The role of honey in the management of wounds. Journal of Wound Care8:415–418.

Ngo QD, Vickery K, Deva AK. 2012. The effect of topical negative pressure on wound biofilmsusing an in vitro wound model. Wound Repair and Regeneration 20:83–90DOI 10.1111/j.1524-475X.2011.00747.x.

Pankuch GA, Jacobs MR, Appelbaum PC. 1998. Postantibiotic effect and postantibiotic sub-MICeffect of quinupristin-dalfopristin against gram-positive and -negative organisms. AntimicrobialAgents and Chemotherapy 42:3028–3031.

Projan SJ, Youngman PJ. 2002. Antimicrobials: new solutions badly needed. Current Opinion inMicrobiology 5:463–465 DOI 10.1016/S1369-5274(02)00364-8.

Ranall MV, Butler MS, Blaskovich MA, Cooper MA. 2012. Resolving biofilm infections: currenttherapy and drug discovery strategies. Current Drug Targets 13:1375–1385DOI 10.2174/138945012803530251.

Sen CK, Gordillo GM, Roy S, Kirsner R, Lambert L, Hunt TK, Gottrup F, Gurtner GC,Longaker MT. 2009. Human skin wounds: a major and snowballing threat to public healthand the economy. Wound Repair and Regeneration 17:763–771DOI 10.1111/j.1524-475X.2009.00543.x.

Sherlock O, Dolan A, Athman R, Power A, Gethin G, Cowman S, Humphreys H. 2010.Comparison of the antimicrobial activity of Ulmo honey from Chile and Manuka honey againstmethicillin-resistant Staphylococcus aureus, Escherichia coli and Pseudomonas aeruginosa. BMCComplementary and Alternative Medicine 10:47 DOI 10.1186/1472-6882-10-47.

Simon A, Traynor K, Santos K, Blaser G, Bode U, Molan P. 2009. Medical honey for woundcare–still the ‘latest resort’? Evidence-Based Complementary and Alternative Medicine 6:165–173DOI 10.1093/ecam/nem175.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 24/25

Stepanovic S, Vukovic D, Dakic I, Savic B, Svabic-Vlahovic M. 2000. A modified microtiter-platetest for quantification of staphylococcal biofilm formation. Journal of Microbiological Methods40:175–179 DOI 10.1016/S0167-7012(00)00122-6.

Stephens JM, Schlothauer RC, Morris BD, Yang D, Fearnley L, Greenwood DR, Loomes KM.2009. Phenolic compounds and methylglyoxal in some New Zealand Manuka and Kanukahoneys. Food Chemistry 120:78–86 DOI 10.1016/j.foodchem.2009.09.074.

Stocks SM. 2004. Mechanism and use of the commercially available viability stain, BacLight.Cytometry Part A 61:189–195 DOI 10.1002/cyto.a.20069.

Subrt N, Mesak LR, Davies J. 2011. Modulation of virulence gene expression by cell wall activeantibiotics in Staphylococcus aureus. Journal of Antimicrobial Chemotherapy 66:979–984DOI 10.1093/jac/dkr043.

Weston RJ, Brocklebank LK, Lu Y. 2000. Identification and quantitative levels ofantibacterial components of some New Zealand honeys. Food Chemistry 70:427–435DOI 10.1016/S0308-8146(00)00127-8.

Wolcott RD, Rhoads DD, Dowd SE. 2008. Biofilms and chronic wound inflammation. Journal ofWound Care 17:333–341.

Zmantar T, Kouidhi B, Miladi H, Mahdouani K, Bakhrouf A. 2010. A microtiter plate assayfor Staphylococcus aureus biofilm quantification at various pH levels and hydrogen peroxidesupplementation. New Microbiologica 33:137–145.

Lu et al. (2014), PeerJ, DOI 10.7717/peerj.326 25/25